Platelets are anuclear cell fragments shed from bone marrow megakaryocytes that circulate through the vascular system. Platelets play two major roles in hemostasis: adhesion to the walls of damaged blood vessels, leading to the formation of a platelet plug; and activation of platelet-dependent blood coagulation pathways, essential for the formation of a fibrin clot. Thus platelets are critical for blood clot initiation and the control of patient bleeding (i.e., hemorrhaging) due to injury, trauma, bleeding disorders, or medical intervensions.
Blood clot initiation begins with the adhesion of platelets to the wall of an injured vessel. von Willebrand factor (“vWF”) bind to platelets and help them attach to collagen present in the surrounding tissues at the site of injury forming the initial platelet clot. Fibrinogen, a soluble plasma protein, is converted to insoluble strands of fibrin by the enzyme thrombin (which is activated by activated Factor X). Thrombin also converts Factor XIII to activated Factor XIIIa. Fibrin filaments are then cross-linked by Factor XIIIa to form a fibrin-platelet mesh referred to as a fibrin clot.
Coagulation factor X can be activated by two distinct pathways, termed the extrinsic and intrinsic pathways. The extrinsic pathway is activated by sub-endothelial tissue factor (“TF”), which is not normally present in the lumen of intact blood vessels. Blood vessel disruption exposes circulating clotting factors to TF. Factor VII is activated to Factor VIIa, which in turn activates Factor X to Factor Xa. In contrast, activation of the intrinsic pathway does not always require TF, especially in vitro. The intrinsic pathway is activated by contact with negatively charged substances, such as contact with sub-endothelial collagen in vivo, or glass in vitro—Factor XII is activated to Factor XIIa, which converts Factor XI to Factor XIa. In the presence of calcium ions, Factor XIa activates Factor IX to IXa, which in turn combines with Factor VIII to activate circulating Factor X. Thus, the activation of blood clotting is dependent on a cascade of several coagulation factors in conjunction with the activity of platelets. In addition, one or more coagulation factor tests may be performed to evaluate the concentration and/or activity of specific coagulation factors (i.e., fibrinogen, vWF, Factor V, Factor VII, Factor VIII, Factor IX, Factor XI, and Factor XII).
As such, patient hemostasis is often evaluated by analyzing a patient blood sample for platelet counts, various markers involved in blood clotting, and clot forming ability, including the amount of time it takes to clot initiation. The coagulation assays measuring the prothrombin time (“PT”) and the partial thromboplastin time (“PTT”) or otherwise known as activated partial thromboplastin time (“aPTT”) are used to evaluate the extrinsic and intrinsic coagulation pathways, respectively. Thromboelastography (“TEG”) is another method of testing the efficiency of blood coagulation. It is mainly used in surgery and anesthesiology and has been established as a sensitive test for hemostatic function in several clinical settings. TEG measures coagulation factor activity, platelet function, clot strength, and fibrinolysis by triggering clot formation followed by computerized coagulation analysis. These assays allow the detection of blood clotting diseases, disorders, or complications arising from diseases, treatments, injuries, surgical procedures, and/or trauma.
While excessive clotting can cause blood vessel occlusion, insufficient or weak clotting can result in excessive blood loss. Treatment of platelet deficiency (a condition referred to as thrombocytopenia) involves the transfusion of stored, concentrated, platelets prepared from donated blood units. Refrigeration of platelets prior to transfusion inhibits platelet function and results in rapid clearance from the circulation (Murphy et al., “Effect of Storage Temperature on Maintenance of Platelet Viability—Deleterious Effect of Refrigerated Storage,” N Engl. J. Med. 280:1094-1098 (1969)). Thus, platelets are best stored at room temperature. Because room temperature storage has the downside of promoting bacterial growth, platelet storage is ordinarily limited to a maximum of 5 days, making platelet inventory management extremely challenging (Kaufman R M, “Platelets: Testing, Dosing, and the Storage Lesion—Recent Advances,” Hematology Am. Soc. Hematol. Educ. Program. 1:492-6 (2006)).
Stored platelets survive and function after transfusion with varying degrees of success. Platelets that have been gently prepared and then immediately transfused without a significant storage interval (within 24-48 hours of donation) have uniformly high recovery, good survival, and preserved function. However, practically this is not possible since blood products have to be cleared from any infectious diseases before transfusion; a process that take up to 48 hours. Thus, the earliest platelet transfusion is at least 48 hours old. By contrast, after storage for 7 days, occasional platelet units will actually become nonviable, such that these platelets demonstrate virtually no survival following transfusion (Rinder et al., “In Vitro Evaluation of Stored Platelets: Is There Hope for Predicting Posttransfusion Platelet Survival and Function?,” Transfusion. 43:2-6 (2003)).
Nonviability is most often seen following severe pH and metabolic derangement. During storage, platelets continue to be metabolically active. Products of metabolism such as lactate accumulate, and the pH falls. It has been shown that if the pH drops below 6.0-6.2, survival in vivo is severely diminished. Platelet aggregation responses to a number of agonists also significantly drop during storage (Kaufman R M, “Platelets: Testing, Dosing, and the Storage Lesion—Recent Advances,” Hematology Am. Soc. Hematol. Educ. Program. 1:492-6 (2006)). No in vitro assay has yet been validated to reliably predict platelet survival in vitro (Kaufman R M, “Platelets: Testing, Dosing, and the Storage Lesion—Recent Advances,” Hematology Am. Soc. Hematol. Educ. Program. 1:492-6 (2006)).
Platelets, like other cell types, shed small 0.1-1 micron fragments of their plasma membrane (referred to as microparticles). In particular, platelet microparticles (“PMPs”) account for approximately 70-90% of the total microparticles (“MPs”) in circulation. The average concentration of PMP in circulation in a healthy individual is approximately 2×104 PMP/μl. During storage, the number of PMPs present in a platelet preparation increases with time, such that large quantities of PMPs are transfused together with platelets (Heijnen et al., “Activated Platelets Release Two Types of Membrane Vesicles: Microvesicles by Surface Shedding and Exosomes Derived from Exocytosis of Multivesicular Bodies and Alpha Granules,” Blood. 94:3791-94 (2000)). In vivo, PMPs have been observed in clinical situations associated with platelet activation and procoagulant activity (i.e., heparin-induced thrombocytopenia, arterial thrombosis, idiopathic thrombocytopenic purpura, thrombic thrombocytopenia, and sickle cell disease) (Hughes et al., “Morphological Analysis of Microparticle Generation in Heparin-Induced Thrombocytopenia,” Blood. 96:188-94 (2000) and Mallat et al., “Elevated Levels of Shed Membrane Microparticles with Procoagulant Potential in the Peripheral Circulating Blood of Patients with Acute Coronary Syndromes,” Circulation. 101:841-3 (2000)), suggesting roles for PMPs in coagulation, cellular signaling, vascular injury, and homeostasis (Hargett et al., “On the Origin of Microparticles: From ‘Platelet Dust’ to Mediators of Intercellular Communication,” Pulm. Circ. 3:329-40 (2013)).
Methods for the isolation of platelet microparticles have been previously described in the literature. For example, international patent publication WO 1990/012581 to Chao discloses a method for the generation of PMPs from stored blood. U.S. Pat. App. Publ. 2006/0034809 to Ho et al. discloses compositions comprising freeze-dried platelets and freeze-dried platelet microparticles. Furthermore, U.S. Pat. No. 8,105,632 to Jy et al. discloses three alternative methods for PMP production. The methods include acidification of stored platelet concentrates, the sonication of freshly isolated or stored platelets, and treatment of platelets with calcium ionophore in the presence of calcium to induce platelet activation and PMP production.
Although numerous groups have developed methods for making isolated platelet microparticle compositions, there exists a need for improved PMP compositions that can restore normal hemostasis in bleeding patients, that are freshly prepared, suitable for long-term storage (up to 12 months), and that remain efficacious for periods extending beyond 5 days from preparation.
The disclosure herein is directed to overcoming these and other deficiencies in the art.